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Abstract

Background

The isolation of green fluorescent protein (GFP) and the development of spectral variants
over the past decade have begun to reveal the dynamic nature of protein trafficking
and organelle motility. In planta analyses of this dynamic process have typically been limited to only two organelles
or proteins at a time in only a few cell types.

Results

We generated a transgenic Arabidopsis plant that contains four spectrally different
fluorescent proteins. Nuclei, plastids, mitochondria and plasma membranes were genetically
tagged with cyan, red, yellow and green fluorescent proteins, respectively. In addition,
methods to track nuclei, mitochondria and chloroplasts and quantify the interaction
between these organelles at a submicron resolution were developed. These analyzes
revealed that N-ethylmaleimide disrupts nuclear-mitochondrial but not nuclear-plastids
interactions in root epidermal cells of live Arabidopsis seedlings.

Conclusion

We developed a tool and associated methods for analyzing the complex dynamic of organelle-organelle
interactions in real time in planta. Homozygous transgenic Arabidopsis (Kaleidocell) is available through Arabidopsis
Biological Resource Center.

Background

Advances in fluorescence microscopy [1] and the development of multiple spectral variants of the green fluorescent protein
(GFP) [2] have revolutionized plant cell biology. These technological developments have produced
a wealth of published information related to protein localization, and are currently
being exploited to analyze protein-protein interactions [3]. Recently, this fluorescence technology has begun to be applied to an analysis of
the dynamic motility of several organelles, including mitochondria [4-6] nuclei [7] and Golgi [8]. Much of this information has been deposited in the plant organelle database (PODB)
[9]. However, the majority of these plant studies have been performed using transient
assays or have involved marking only a few organelles with one or two fluorescent
proteins, making it difficult to perform detailed analyses of the interaction or association
between multiple organelles. In both plants and animals, some organelles are localized
in close proximity to one another such that the distance that separates them is shorter
than the resolution limit of a conventional fluorescence microscope (< 300 nm), but
longer than the resolution limit of a conventional electron microscope (> 20 nm).
In the context of the studies presented here, we describe these close localizations
of organelles as organelle-organelle interactions (OOIs).

A classical example of OOIs in plants are those involving chloroplasts, peroxisomes
and mitochondria in green tobacco leaves, observed by electron microscopy [10]. Because enzymes required for photorespiration are stored separately in these organelles,
secondary metabolites, such as glycolate and glycine, must be transported among these
organelles. Hence, in this case, the OOIs may facilitate photorespiration by maintaining
short transportation distances. Using confocal microscopy, physical interactions between
the nucleus and plastids were recently described in tobacco hypocotyl epidermal cells
[11], and signal transduction between the nucleus and plastids has also been reported
[12]. Kwok and Hanson have hypothesized that nucleus-plastid signaling may be facilitated
by OOIs that reduce the distance between the nucleus and plastids [11]. In animal cells, OOIs between the nucleus and mitochondria have also been described
[13]. In a study of ATP transport, physical interactions between nuclei and mitochondria
were observed in cultured cells using both confocal and electron microscopy. Although
nuclei are not capable of producing ATP, a large amount of it is required for nucleus-specific
functions, such as transcription and mRNA transport. Thus, nucleus-mitochondria OOIs
may facilitate ATP transport from the mitochondria to the nucleus [13].

Although biologically important roles for OOIs have been hypothesized as described
above, the nature of organelle-organelle physical interactions is poorly understood.
Indeed, molecules that mediate the physical interactions between nuclei and plastids
or mitochondria have yet to be identified. Organelles may interact with each other
through an active molecular recognition mechanism or perhaps, if cellular space is
limited, through a passive process. In this study, we sought to determine which phenomenon
– active or passive recognition – is correct by addressing the following questions:

1. How many plastids and mitochondria are in each cell?

2. How much space is available in a cell for the nucleus, plastids and mitochondria?

3. Where do these organelles locate in the three-dimensional (3-D) space of the cell?

4. How long do the OOIs last?

5. Are specific molecular interactions involved in mediating OOIs?

To begin to answer these questions, we have developed a transgenic Arabidopsis plant
as a tool for analyzing the dynamics of multiple organelles and multiple OOIs in real
time in planta.

Figure 1.Transgene constructs and their expression in the Kaleidocell line. (A) The fusion genes were inserted downstream of the 35S cauliflower mosaic virus promoter
(35SP). Expression cassettes of Gal4-CFP and CoxIV-YFP were inserted in tandem in
the transgene region [between the right (RB) and left borders (LB)]. Expression cassettes
of Cam53BD-GFP and RecA-RFP were inserted in the transgene region. Unique sites digested
by BamHI in the transgenes and the predicted sizes (kbp: kilo base pairs) between the BamHI sites and to the RB were shown on the top of the constructs. The size of the 35SP
is also shown on the bottom of the construct. A transgenic plant expressing Gal4-CFP
and CoxIV-YFP was crossed with a transgenic plant expressing Cam53BD-GFP. The resulting
plant was further crossed with a transgenic plant expressing RecA-RFP. The final transgenic
plant expressing all four transgenes was designated as the Kaleidocell line. Gal4:
Saccharomyces cerevisiae nuclear protein, CoxIV: Saccharomyces cerevisiae cytochrome oxidase IV, Cam53BD: petunia calmodulin CaM53 binding domain, RecA: Arabidopsis DNA repair protein. (B) Southern blotting analysis of the BamHI digested genomic DNA of homozygous Kaleidocell. Lane M: 1 kb DNA Ladder (NEB),
Lane BamHI: BamHI digested genomic DNA. The sizes of the DNA ladder are shown on the left. The DNA
probe is shown on the right bottom. (C) Fluorescence microscope images of a protoplast from the Kaleidocell line. The protoplast
was observed with four different fluorescence filter sets: cyan, green, yellow, or
red. Captured images were merged to generate a single image of the protoplast. Scale
bars = 10 μm.

To confirm transgene integration and to predict transgene copy numbers, we performed
Southern blots on BamHI-digested genomic DNA from homozygous Kaleidocell using a probe
encoding the promoter region of each of these transgenes (Figure 1B). We detected four bands, indicating that a single copy of each expression cassette
(CFP, RFP, YFP, and GFP) was integrated in the Kaleidocell genome. One of the bands
appeared at approximately 1.4 kbp, which is 2.0 kbp shorter than expected-shortest
size (3.4 kbp). The reduced size of this band suggests that one of the T-DNAs did
not integrate completely into the genome. However, the fact that all four spectral
variants were visible in Kaleidocell specimens indicates that this truncation of T-DNA
had no apparent effect on the functionality of the four transgenes.

To confirm proper accumulation of each fluorescent protein, protoplasts were generated
from one-week-old homozygous Kaleidocell seedlings (Figure 1C). Within a single protoplast, we detected cyan, red, yellow, and green fluorescence
in the nucleus, plastids, mitochondria, and plasma membrane, respectively. The fluorescence
spectrums of CFP and GFP overlap with one another, as do those of GFP and YFP. Accordingly,
it can be challenging to distinguish CFP, GFP and YFP signals in a single cell [21]. Although these signals can be separated through the use of spectral imaging and
linear un-mixing of image data [22], the microscopes and software used in this study did not allow us to do so. Furthermore,
the petunia calmodulin CaM53 that tags plasma membranes tended to accumulate in nuclei
by blocking isoprenoid biosynthesis or depleting carbons [19]. To distinguish organelles from one another in this study, we used a combination
of fluorescent signals and organelle size. The nucleus was a clearly recognizable
entity with a diameter larger than 5 μm. Meanwhile, plastids were entities with diameters
just larger than 2.5 μm, and mitochondria were the smallest recognizable objects with
a diameter greater than 0.7 μm.

The fluorescent proteins in Kaleidocell seedlings (3 days to 2 weeks post-germination)
were easily detected in the hypocotyls and roots of all individuals analyzed (Figure
2A and 2B). However, three weeks post-germination, the fluorescent protein signals were not
as readily detected. Although the reason for this time-dependent reduction in signal
intensity is unknown and requires further study, it is likely that the transgene loci
may be affected by local chromatin modifications, which alter transgene expression
levels at distinct stages of plant development.

Figure 2.Expression of the transgenes in seedlings in the Kaleidocell line. (A) Fluorescence micrograph of a root meristem from the 7 days-old Kaleidocell line. The
root meristem was observed with three different fluorescence filter sets: cyan, yellow,
and red. Captured images were merged to generate a single image. (B) Fluorescence micrograph of epidermal cells from the cotyledon of a 7 day-old Kaleidocell
plant. The cotyledon was observed with three different fluorescence filter sets: cyan,
yellow, and red. Captured images were merged to generate a single image. Scale bars
= 50 μm.

Because Kaleidocell is homozygous for all four fluorescent proteins, plants can be
propagated without using a selection reagent such as kanamycin. Additionally, we have
found that the fluorescence of all four fluorescent proteins remains stable in subsequent
generations (data not shown).

Mounting living seedlings on a microscope stage

Living plants can be adhered to a coverglass for a number of hours using a gelatin-
or agar-based gel mounting medium [23]. Although this method generally works well, it is not suitable for high-resolution
3-D microscopy because of optical scattering and aberrations caused by the gel (Kato
unpublished). To optimize the quality of the signal captured from the Kaleidocell
plant, we developed a simple method that allowed us to make extended observations
(hours) of whole seedlings with minimum optical scattering or other aberrations. Our
method uses water-soaked Kimwipes to provide 1) a mounting medium (water) that fills
the empty space between the sample and the coverglass and reduces dehydration of the
sample being analyzed, and 2) a force that gently pushes the seedling towards the
coverglass (Figure 3A–C). During our experiments, Arabidopsis seedlings mounted in this manner grew for over 12 hours on a microscope stage (data
not shown).

Figure 3.Mounting a seedling on an inverted fluorescence microscope. (A) Preparation: each seedling was laid on a chamber-coverglass, and layers of Kimwipes
were folded and soaked with distilled water. (B) Mounting: water-soaked Kimwipes were wrung out onto the chamber-coverglass. A bottom
view of the chamber-coverglass is shown. (C) Microscope stage setting: a microscope slideglass was used as a lid for the chamber-glass.
The chamber-glass was fixed on the stage with labeling tape. (D) Alternative mounting method. Scale bars = 4 cm.

In addition, a convenient method also was developed to enclose Kaleidocell in a microscope
slide (Figure 3D). After germinating seeds for 3 days on water-soaked blotting paper in a plastic
Petri dish, a microscope slide with a one-well adhesive spacer (20 mm in diameter,
0.12 mm deep) was used to mount Kaleidocell with water. A glass cover slip was used
to seal a seedling in this chamber. Although it may be difficult to continuously observe
single cells for extended periods due to the gradual movement of Kaleidocell in water,
it is possible to obtain sequential high-resolution images over a short period of
time (e.g., 10 minutes). This method may be useful to quickly test or demonstrate
the feasibility of Kaleidocell.

Detection of nuclei, plastids and mitochondria in epidermal and cortical cells of
the Kaleidocell root

To visualize the multiple fluorescent proteins marking the different organelles in
Kaleidocell, we obtained a stacked confocal scanning microscopic image of a Kaleidocell
seedling root (Figure 4). The resolution of the objective lens used in this scan was calculated at 0.25 μm
in the x- and y-axes and 0.65 μm in the z-axis. Because of the limitations of our
laser lines, we acquired GFP (plasma membranes) and YFP (mitochondria) signals in
the same channel. However, as noted above, the size and pattern of each of these organelles
made it easy to distinguish the plasma membrane from the mitochondria.

Figure 4.Detection of nuclei, plastids, and mitochondria in a Kaleidocell root. (A) A maximum projection image of a vertical series of scanning confocal micrographs of
a root of a Kaleidocell seedling. The root grew along the x-axis. A total of 17 epidermal
and 8 cortical cells (green lines) were identified. Cell boarders at the ends of the
x-axis (top and bottom of the image) were beyond the scanning area. Within a total
of the 25 cells, 11 nuclei (blue spots), 255 plastids (red spots), and 10,158 mitochondria
(green spots) were identified. A white thin horizontal line in the middle of the image
indicates a point where a cut-through image was generated. (B) Cut-through image of the stacked image. Image contrast was adjusted to enable clear
visualization of the cell borders. Scale bars = 10 μm.

A total of 25 cells, 17 epidermal and eight cortical, were clearly identified within
a 230.2 × 230.3 × 30.9 μm scan area (Figure 4). Of the 25 identified cells, 11 nuclei were inside the scan area during acquisition.
The number of plastids and mitochondria were determined using imaging software. Point-like
structures (spots) were automatically identified, and 255 RFP spots (diameter > 2.5
μm) and 10,158 GFP/YFP spots (diameter > 0.7 μm) were detected. Hence, we estimated
that a single cell in the root of the Kaleidocell seedling contained approximately
10 plastids and 400 mitochondria. Mitochondria do not exist in cells as discrete units
and could be smaller than our microscopic resolution [4,5], so the actual number of mitochondria may be higher or lower than our reported numbers.
However, the number suggested by our analysis is similar to that predicted by the
amount of mitochondrial DNA in animal cells [24], providing a measure of confidence that the number of mitochondria we were able to
visualize in a single cell would be close to the actual number.

Three-dimensional positioning of nuclei, plastids and mitochondria

The stacked confocal scanning microscopic image was converted to a 3-D model and visually
inspected using imaging software to determine the location of organelles in space
(Figure 5 and see Additional file 1). We found that almost all organelles were located in the cortical region of the
cells. By manually identifying cellular locations of nuclei, plastids and mitochondria,
we estimated that 60 ± 12.5% (n = 10) of the cell volume did not contain these organelles.
This suggests that the cytosolic area in which nuclei, plastids and mitochondria can
move comprises approximately 40% of the total cellular volume in epidermal and cortical
cells of Arabidopsis roots. However, it has been demonstrated previously that expanded vacuoles in root
cells [25] push nuclei, plastids and mitochondria to the cortex. Thus, the total cellular volume
available to these organelles may vary as a function of vacuole size and/or complexity
in different cells or cell types.

Additional file 1.A movie file of Figure 5. A Quick time movie file of Figure 5. A point of view is simultaneously changed in the virtual three-dimensional world.

Figure 5.Localization of nuclei, plastids, and mitochondria in a 3-D model. The stacked image was converted to a 3-D model (blends projection). In this projection,
viewing directions and their transparencies were blended to produce a 3-D perspective
of the structure. Viewpoints of each image (A to D) are indicated on the bottom. The
box indicates the 3-D coordination of the stacked image. The different colors on each
wall indicate the different dimensions. Arrows with a letter indicate the viewpoints
of each image. A black ellipse on the bottom of the box indicates the position of
a nucleus. (A) The blends projection of the stacked image of Figure 4(A). A white rectangle indicates
the area enlarged in (B, C, and D). (B) Top view. A nucleus (blue) interacts with plastids (red) and mitochondria (green).
The 3-D coordination was not clear in this viewpoint. (C) Side view of the same nucleus. Note that the plastids lay on the nucleus. Also note
that the majority of the organelles localize to the cortex of the cells. (D) Bottom view of the same nucleus. Note that the nucleus lays on the mitochondria. Scale
bars = 10 μm.

Dynamics of nuclear interactions with plastids and mitochondria

To simultaneously investigate the dynamics of OOIs between nuclei and plastids, and
nuclei and mitochondria, we tracked the movements of each organelle in a single root
epidermis for 3 h, at 1 min intervals (Figure 6, Table 1, and see Additional file 2).

Figure 6.Movements of nuclei, plastids, and mitochondria. Stacked deconvolution micrographs of root epidermal cells during a 3 h and 40 min
long time-lapse observation. (A) Image captured at 2 h and 46 min into the observation period. Nuclei, plastids, and
mitochondria appear blue, red, and green, respectively. Triangles indicate the nuclei.
Scale bar = 10 μm. (B) Movements of nuclei tracked from 2 h and 27 min to 3 h and 40 min (a total tracking
duration of 103 min). Each track is displayed as a line. Line color indicates the
time point, corresponding to the time color bar on the right bottom. Solid-line circles
indicate areas where the nuclei moved. Dashed-line circles indicate an area where
the nucleus stalled. (C) Movements of plastids. Movements are indicated by solid- and dashed-line circles as
in (B). (D) Movements of mitochondria. Movements are indicated by solid- and dashed-line circles
as in (B). Notice that the tracks of the plastids and mitochondria are similar to
that of the nuclei.

During this time, we observed nuclear movements similar to those reported previously
[7]. For example, one nucleus moved constantly at an average speed of 2.4 μm/min for
70 min covering a distance of 122 μm; another nucleus within the same tissue did not
move as far (4-μm travel distance over 70 min, Figure 6 and see Additional file 3). A previous study indicated that nuclear movements in Arabidopsis root hairs correlated with cell growth [26]. Hence, the growth status of individual cells in the root epidermis may be reflected
in nuclear movements.

Plastid movements have also been previously reported to vary [6]. We found the plastids that interacted with nuclei moved with the nuclei over the
3-h observation interval (Figure 6 and see Additional file 4), indicating that the nucleus constantly interacts with the same population of plastids,
independent of their movement within the cell.

The mitochondria were smaller and moved faster than the other organelles, making them
difficult to track individually. The software we used was unable to track the movement
of the mitochondria for an extended period (Table 1). However, we did observe that the mitochondria that interacted with and surrounded
the nuclei moved in the same direction as the nuclei (Figure 6 and see Additional file 5). Although this does not conclusively demonstrate that the nucleus was interacting
with the same population of mitochondria, it does suggest that the nucleus is constantly
interacting with mitochondria.

Pharmacological analysis of OOIs between nuclei and plastids or mitochondria

Unlike most other eukaryotes, actin microfilaments control organelle movements in
higher plants (reviewed in [27,28]). Actin microfilaments, together with myosin motor proteins, also control cytoplasmic
streaming [27]. It is also known that microtubules, which control organelle movements in other eukaryotes,
contribute to the positioning of organelles in higher plants[28].

To explore the nature of the nucleus-plastid and nucleus-mitochondria OOIs, we performed
a pharmacological screen to search for factors that might alter these OOIs. We exposed
Kaleidocell seedlings to five different pharmacological compounds that have been used
previously to study the positions and movements of nuclei, mitochondria or plastids
in higher plants [7,29-31]. The first compound, Latrunculin B, decreases the amount of polymerized actin and
immobilizes nuclei [7] and mitochondria [29]; it also disrupts plastid positioning [30]. The second compound, oryzalin, influences tubulin polymerization and disrupts the
positioning of mitochondria [29]. The third compound, 2,4-dinitrophenol (DNP), accelerates depletion of ATP from the
cells and immobilizes mitochondria [29]. The fourth compound, 2-monoxime (BDM), is a mammalian myosin ATPase inhibitor that
immobilizes mitochondria and also causes immobilized mitochondria to aggregate [29]. The fifth compound, N-ethylmaleimide (NEM), causes alkylation of proteins and immobilizes
nuclei [7], mitochondria [29] and plastids [31], and also causes immobilized mitochondria to aggregate [29].

Nucleus-mitochondria interactions and nucleus-plastid interactions were manually identified
under a fluorescence microscope using size, morphology and spectral fluorescence to
identify each type of organelle (Figure 7). In the absence of pharmacological compound exposure, 71% of nuclei were observed
to interact with mitochondria (n = 128, 6 independent experiments). This percentage
was lower than that obtained using confocal imaging (~100% of the observed nuclei),
suggesting that the lower resolution of the objective lens used on the epifluorescence
microscope may limit the ability to identify all nucleus-mitochondria OOIs. Alternatively,
the inability to acquire 3-D images may have been an issue, resulting in mitochondria
located beneath nuclei being missed during OOI detections. However, this approach
was capable of detecting a majority of nucleus-mitochondria OOIs and the quantitative
differences did not interfere with our ability to analyze the effects of pharmacological
compounds on these OOIs.

To the extent that molecules involved in nucleus-plastid and/or nucleus-mitochondria
OOIs are affected by a given pharmacological compound, the prediction is that fewer
OOIs would be observed between these organelles in cells treated with the corresponding
compound. No significant change in the percentage of nucleus-mitochondria OOIs was
observed in seedlings treated with latrunculin or oryzalin compared to mock-treated
seedlings (Figure 7A). These results suggest that cytoskeletal proteins, such as actin and microtubules,
do not participate in this type of OOI. In contrast, NEM significantly reduced the
percentage of nucleus-mitochondria OOIs (0.3-fold, P ≤ 0.01), suggesting that molecules
sensitive to NEM-mediated alkylation may be involved. Moreover, dual exposure to NEM
and latrunculin or NEM and oryzalin did not induce a further reduction in the interaction
rate beyond that of NEM alone, indicating the absence of a synergistic effect. It
is impossible to predict which molecules might control the nucleus-mitochondria OOIs
on the basis of this experiment because NEM immobilizes many other organelles and
impacts fluid dynamics (cytoplasmic streaming) in plant cells [31]. A more extensive analysis, perhaps involving random mutagenesis of the Kaleidocell
line, will be required to identify the molecules involved in nucleus-mitochondria
OOIs. Analyses of nucleus-plastid OOIs in these same experiments did not reveal any
changes in the presence of any of these pharmacological compounds (Figure 7B). Although these results do not allow us to form any conclusions about the molecular
aspects of nucleus-plastid OOIs, they do suggest that molecules or mechanisms other
than those involved in nucleus-mitochondria OOIs are responsible for regulating nucleus-plastid
OOIs. The difference between nucleus-mitochondria and nucleus-plastid interactions
also suggest that these interactions are active interactions and not passive interactions.

Conclusion

We demonstrate that Kaleidocell is a useful tool to begin the exploration of OOIs.
We have identified nucleus-chloroplast and nucleus-mitochondria interactions in root
epidermal cells, and have demonstrated that NEM disrupts nucleus-mitochondria OOIs
but not nucleus-plastid OOIs. These results suggest that different mechanisms are
involved in regulating nucleus-mitochondria and nucleus-plastid OOIs. Kaleidocell
should also be useful in experiments designed to determine if cell type, developmental
stage and/or exogenous stimuli are capable of regulating or altering these interactions.
The homozygous Kaleidocell line is available through Arabidopsis Biological Resource
Center with the stock number CS16303.

Methods

Fluorescent proteins to tag mitochondria and nuclei

The mitochondrial localization signal sequence in cytochrome oxidase IV (CoxIV) of
Saccharomyces cerevisiae was amplified from pCK CoxIV-GFP [17] and fused to yellow fluorescent protein EYFP (Clontech, CA) by a three step PCR amplification.
The primers used for amplifying the coxIV signal peptide were LM128 (5'-AgggATCCAAAATggTTTCACTACgTCAATCTATAAgA-3')
and LM129 (5'-TTgCTCACCATgggTTTTTgCTgAAgCAgATATCT-3'). The primers used for amplifying
EYFP from the pEYFP-C1 (Clontech) were LM130 (5'-AgCAAAAACCCATggTgAgCAAgggCgAggAgCTgT-3')
and LM131 (5'-ACgAgCTCACTTgTACAgCTCgTCCATgCCgA-3'). The products of these two reactions
were combined and a third amplification was performed using LM128 and LM131. The product
of this amplification was cloned into pGEM-T (Promega) and sequenced. The coxIV::EYFP
was subsequently removed from this vector using the BamHI and SacI sites that were
included in primers LM128 and LM131, respectively. The coxIV-EYFP BamHI/SacI fragment
was then inserted into pBI121 (Clontech) at BamHI/SacI, thereby replacing the β-glucuronidase
gene of the pBI121 vector (Clontech), forming pBI121-CoxIV::EYFP. A HindIII and EcoRI
digested fragment that encoding 35SP (cauliflower mosaic virus 35S promoter)-CoxIV::EYFP-NosT(nopalin
synthase terminator) in pBI121-CoxIV::EYFP was inserted in HindIII and Sse8387I digested
pKW102 vector via DNA adapter that encodes EcoRI and PstI. The HindIII and Sse8387I
sites are located in an upstream of 35SP in pKW102 that expresses Gal4::ECFP::NLS,
a chimeric gene encoding yeast nuclear localized protein Gal4, a nuclear localization
signal from SV40 large T-antigen, and the cyan fluorescent protein ECFP (Clontech).
The resulting vector, designated as pKW102-CY-MODC, hence expresses both CoxIV::EYFP
and Gal4::ECFP::NLS from a single T-DNA.

Fluorescent proteins to tag plastids

The EYFP region of pEYFP-C1 vector (Clontech) was removed with NheI/SacI and subcloned
into pBluescript SK+ (Stratagene) at XbaI/SacI to form pBSK+-EYFP. This vector was
subsequently digested with BamHI/SacI and the EYFP containing region was subcloned
into pBI221 (Clontech) digested with the same restriction enzymes, forming the pBI221-EYFP
vector. A 300 bp fragment, corresponding to the signal peptide of the Arabidopsis
RecA protein that localized in chloroplasts (NM_106556) was PCR amplified from an
Arabidopsis cDNA library using primers LM116 (5' CgggATCCATggATTCACAgCTAgTCTTgTC
3') and LM117 (5' gAAgATCTTCCATAgCTgCCTCTAAAgCCTT 3'). This fragment was cloned into
the pGEM-T vector (Promega) and sequenced. The RecA signal peptide was subsequently
removed from this vector using the restriction sites that were incorporated in the
primers (BamHI/BglII) and subcloned into the BamHI site of pBI221-EYFP forming the
pBI221-RecA::EYFP vector. The EYFP region of this vector was replaced with the red
fluorescent protein DsRed2 (Clontech) at the AgeI/SacI sites, forming the pBI221-RecA::DsRed2
vector. The region containing RecA::DsRed2 was then transferred to pEL103 [21] at the BamHI/SacI sites, forming the pEL103-RecA::DsRed2 vector.

Plant transformation

Agrobacterium tumefaciens strain GV3101/pMP90 was transformed with the vectors, pKW102-CY-MODC, pEL103-RecA::DsRed2,
and pEL103-GR respectively. These Agrobacterium strains were then used to transform Arabidopsis thaliana (Col-0) as described previously [32].

Plant screening and crossing

Zygosities of each transgenic line were determined by their sensitivity to kanamycin
selection and by detection of the fluorescent proteins in their seedlings. Homozygous
lines of transgenic Arabidopsis expressing Gal4-CFP and CoxIV-YFP were crossed to homozygous lines of transgenic Arabidopsis expressing Cam53BD-GFP. The resulting homozygous lines generated from the self pollination
of this cross were then crossed to RecA-RFP homozygous plants to obtain the Kaleidocell
lines. The line that most strongly expressed all transgenes was designated Kaleidocell.17.
Kaleidocell.17 was self-pollinated in the following three generations. The line that
expressed all transgenes in its all progenies was identified and designated Kaleidocell.17.1.7.
The progenies of Kaleidocell.17.1.7 are referred to as Kaleidocell in this report.

Plant growth conditions to establish the Kaleidocell line

The seeds were germinated on 0.75% agar plates containing a half-strength (1/2) MS
salt (Sigma, CA) containing 100 mg/L of kanamycin after sterilizing in 50% diluted
Crolox (Crolox, CA). The plates were kept in an environmental chamber in which a temperature
was set at 21°C and a day cycle was set at 16 hours light and 8 hours dark. The plants
selected were transplanted in soil and grown in the same chamber.

Southern blotting

The Southern blotting was conducted based on the previously published method [15]. Hydroponically cultured homozygous Kaleidocell was used to extract the genomic DNA.

Protoplast preparation

The protoplasts were prepared based on the previously published protocol [33]. About 50 seedlings of the one-week-old Kaleidocell line were used as starting material.

Seedling preparation for the extended-hour observation

An one-weeks-old seedling was transferred to a chambered #1.5 cover glass (Lab-Tek
II, Catalogue number 155360, Nalge Nunc International, IL) and 4 to 5 sheets of 11
× 21 cm water-soaked Kimwipes® (Kimberky-Clark, GA) were folded and piled on the top of the seedling in the chamber
so that a height of the piled Kimwipes were slightly (i.e., ~5 mm) higher than that
of the chamber. A slide (75 mm × 22 mm, Catalogue number 2948, Corning, NY) was used
as a lid for the chamber. The slide-covered chamber was then fixed on a microscope
stage using labeling tape (1.9 mm width, Catalogue number 15-959, Fisher Scientific,
PA) so that the slide could push the water-soaked Kimwipes towards the seedling being
analyzed.

Sample preparation for the pharmacological analysis

One-week-old seedlings were transferred to a 60 × 15 mm Petri dish filled with 10
ml of distilled water. Dimethylsulfoxide (DMSO) or water solution containing pharmacological
compounds was added into the Petri dish and gently shaken for 1 h at about 15 rpm
on a rotory shaker. The seedlings were first transferred to a new Petri dish filled
with distilled water to rinse the compounds and then transferred to a slide (75 mm
× 22 mm) with a 0.12 mm deep spacer (Catalogue number S24736, Invitrogen, CA). Distilled
water was added in the space formed by the spacer and this was sealed with a #1.5
cover glass (Catalogue number 2640, Corning, NY).

The stacked images acquired with the laser scanning confocal microscope were first
deconvolved with the blind deconvolution algorithm (AutoQuant Imaging, NY). The output
levels in each channel were manually adjusted to increase signal-to-background ratios.
The movies of three-dimensional models of Kaleidocell were created using Imaris 5.7.0
(Bitplane AG, Switzerland).

Image analysis: Creating time-lapse movies

The software softWoRx equipped with the Delta Vision® RT Restoration Imaging System was used to capture sequential images for time-lapse
movies. This image data was then used on Imaris 5.7.0 to track organelle movement.

The software ImageJ1.3.7 was used to compile the protoplast images and manually identify
the organelle-organelle interactions. These images were manually manipulated to enhance
the signal-to-background ratio.

Competing interests

The authors declare that they have no competing interests.

Authors' contributions

NK designed the experiments, constructed the DNA vectors, generated and molecularly
analyzed the transgenic plants, and analyzed the 3-D images. DR conducted the pharmacological
analysis and collected the homozygous line of the transgenic plant for donation. MLB
analyzed a root meristem and cotyledon. MB established the homozygous line of the
transgenic plant. YF analyzed the protoplasts. AM and LAM designed and constructed
the RecA-RFP and CoxIV-YFP. All authors read and approved the final manuscript.

Acknowledgements

This publication was made possible in part by NIH Grant Number P20 RR16456 from the
INBRE Program of the National Center for Research Resources. Its contents are solely
the responsibility of the authors and do not necessarily represent the official views
of NIH. NK thanks Dr. Wilhelm Gruissemat at ETH, Zurich for his general gift of the
pGFPnew/BDCaM53wt plasmid, Dr. Eric Lam at Rutgers University, NJ for his support
during the vector construction, Dr. David Burk, and Dr. Tin-Wein Yu at LSU for their
technical supports. The confocal images were obtained during The Tenth Annual 3D Microscopy
of Living Cells Course (University of British Columbia, Canada, June 11 – 27, 2005).
NK thanks the instructors in the course for their technical supports and useful advices.
AM and LM received financial support in part from Fondecyt 1000812, 7000812 and ICM
P02-009-F.